TJ Bliss and Dr. Byron J. Adams, Microbiology and Molecular Biology
Introduction
Nearly every living organism derives benefit from living with one or more species of bacteria during some or all of its lifetime. Numerous studies have described the phenotypic benefit that each or either partner receives from associations of this kind, but the molecular mechanisms of such interactions have not been well described. Understanding beneficial bacterial interactions with their hosts are of interest in that information obtained will have implications in identifying mechanistic components of those interactions that may operate similarly in pathogens and can be exploited in combating infection or in describing the interactions between bacteria and more complex organisms in a community.
The nematode Heterorhabditis bacteriophora (Hb) is one of these model organisms. It has a gut bacterial symbiont, Photorhabdus luminescens (Pl), with which it has so obligately associated that in most cases only the strain of Pl found associated with each subspecies of Hb in the wild is able to support the nematode’s growth and reproduction either in vitro or in vivo1. In vivo, the two are an effective predator of ground-dwelling insects including grubs, weevils and crickets. Hb moves through the soil in search of insect hosts and breaches their outer coverings to regurgitate the entomopathogenic bacteria into the hemolymph of the insect.
I studied the effect that Pl, as a symbiont, exerts on its nematode host and vice versa. Pl exhibits two phenotypic variants, called the primary and secondary phase variants, which show different symbiont-host interactions. Although both are pathogenic when injected into insects, only the primary phase is able to support the growth and development of the nematode. A recent hunt in 30,000 mutants by Joyce and Clarke2 identified a homologue of Escherichia coli’s hexA in Pl that represses the symbiotic phenotype and activates the pathogenic phenotype of Pl in the secondary phase variant. In a similar vein, I wanted to observe whether a screen for a similarly mutated primary phase variant would report any genes in addition to hexA, and analyze the new phenotype for its effect on the in vivo and in vitro growth and development of the nematode.
Research Methods
Mutagenesis. Primary phase cells of spontaneous rifampicin resistant Pl TT01 were subjected to single-gene mutagenesis via conjugation with E. coli S17-1 carrying the suicide tn10 plasmid pLOF which confers kanamycin resistance. After overnight conjugation, 100 ml aliquots of the parent cells were spread onto rif100/Km30 plates. The plates were screened for growth and mutant colonies were isolated. Mutant Screen. The mutants were subjected to 96-hour subcultures in 200 ml of low-osmolarity broth with shaking at 30° C. After 4 subcultures, the mutants were plated onto lipid agar plates, grown for 48 hours and screened for bioluminescence in a dark room. All mutant colonies exhibiting luminescence were isolated and subjected to bioassay. Symbiosis and Pathogenesis Assays. To assay for bacterial ability to symbiotically associate with Hb, axenic eggs were extracted from hermaphroditic nematodes and placed onto 48-hour lawns of mutant cultures and onto of wild type TTO1. The plates were observed for growth and development of monoxenic nematodes. To assay for pathogenicity, dauer juveniles monoxenic with mutant bacteria were used to infect a number of G. mellonella wax worm moth grubs. Phenotype Assays. Mutants were screened for lipase production on SpiritBlue agar. Dye absorption assays were performed on NBTA agar. Motility assays were performed using Motility Indole Ornithine (MIO) agar. Genetics. Bacterial DNA was extracted and PCR-amplified using primers specific to the Pl hexA homologue. To determine if tn10 had inserted itself into the hexA homologue of the mutant, the mutant PCR product was compared to a PCR product of the wild-type amplified hexA homologue.
Results and Conclusions
Of the 1,698 tn10-induced mutants isolated, <100 exhibited any bioluminescent activity after 4 low-osmolarity sub-cultures. Another 4 sub-cultures yielded one constitutively luminescing mutant, m274. The bioluminescence exhibited by m274 after five 96-hour subcultures in low-osmolarity broth was qualitatively greater than that exhibited by wild-type primary phase lawns of Pl TT01. Wild-type cells subjected to the same culturing pressures as m274 exhibited no bioluminescent activity. After 10 days of incubation, most or all of the axenic nematode eggs had hatched on both m274 and on the wild-type bacterial lawn, suggesting that factors needed for symbiosis were active in m274. Dauer juveniles monoxenic with m274 were unable to infect and kill G. mellonella wax worm moth grubs. All 55 insect larvae exposed to dauer juveniles carrying m274 were still alive 5 days after exposure. Thirty-five of 55 insect larvae exposed to juvenile nematodes carrying the wild-type bacteria were dead 3 days post infection. m274 did not produce lipase, appeared bright red on NBTA agar, and was non-motile: all characteristics of secondary phase. PCR amplification and gel comparison show an intact hexA homologue in m274, indicating that some other genetic element must have been disrupted by tn10. The disrupted element could be an upstream or downstream controller of hexA, a gene located upstream of and in the same operon as hexA, or a gene molecularly unconnected to hexA. m274 exhibited many phenotypic characteristics of the secondary phase variant (including inability to kill insects when associated with the nematode host) while still maintaining bioluminescent and symbiotic abilities, suggesting that the knocked-out element does not directly regulate all aspects of phase switching. As mutagenesis did not disrupt the hexA homologue in m274, I concluded that symbiosis and pathogenesis in P. luminescens are regulated by at least one genetic element in addition to the hexA homologue gene. The results of my research were presented at the 2006 Poster Session of the Annual Meeting of the Society of Nematologists in Kauai, Hawaii on June 19, 2006. The poster and my presentation were awarded 1st place in the student poster competition. I acknowledge G. Christopher Bailey, Adler Dillman, Scott Peat, Aaron Smith, Bishwo Adhkari, Dana Blackburn and the other members of the BYU Nematode Evolution Laboratory for assistance and patience. I highly thank my mentor, Dr. Byron J. Adams, for help beyond description throughout the project.